Last
month we looked at some issues regarding cultures and this month I
want to focus on subculturing.

Subculturing,
although it can be time-consuming, is one of the more satisfying
undertakings of the aquatic biologist, once you get the hang of it.

1)
When
To Subculture

This
should always be done at 3:27 a.m.–sorry, but it’s summer
break and I don’t have any students to torment with my bad
jokes right now. Actually, timing subculturing is rather easy to
determine and falls into 2 major categories.

a)
When a culture reaches its peak and the organisms are abundant
and flourishing–this is an ideal time to subculture. Check
your notes and reproduce the conditions of the initial culture as
closely as possible. If it’s an organism which is unusual
and/or of special interest, you may wish to make several subcultures
to make sure that you maintain it in abundance.

b)
When a culture is limping along, but you, nonetheless, have
sufficient organisms to experiment with subculturing, do so. Try
starting a few subcultures by varying the media and keeping careful
records. If one of them succeeds, you can go back to the usual form
of subculturing described above.

2)
Which
Organisms To Subculture

Establish
some clear priorities. Organisms such as Paramecium,
Chilomonas, Halteria, Colpidium, Colpoda, Spirogyra, Vorticella,
Oscillatoria,
etc. are all usually readily available, easily maintained and, as a
consequence, the loss of such cultures is not particularly
problematic, since they are easily re-established. However, if you
have the means, you can have one of your many laboratory assistants
maintain stock culture for you, and lacking such an assistant, you
can probably manage it yourself, since the organisms require
relatively infrequent subculturing. It is a good idea to always have
a stock of Paramecia on hand as it makes a good test organism and
Chilomonas
is
also important since it often makes a good food organism.

I’ll
just mention a few of the kinds of organisms which I have made
efforts to cultivate with varying degrees of success.

a)
First of all, almost any unusual organisms and especially those which
I have not encountered before, I try to culture, be it an algal form,
a protozoan, a lovely aquatic flatworm, a burgundy-colored cyclops, a
strange cladoceran or the bizarrely wonderful Chaoborus
midge larva, that has special little oval-shaped organs which
function like air tanks on a submarine to control its buoyancy. It’s
fascinating to watch them in a jar or small aquarium where they can
hover motionless and then slowly sink or raise themselves toward the
surface. When they find prey, their fierce jaws snap out and seize
the victim faster than you can see.

b)
The large ciliates and a number of the mid-sized ones are especially
intriguing to me.

b1)
Stentor
coeruleus,
the large blue Stentor
is
fairly readily cultured and is a magnificent creature to observe. In
older neglected cultures, it tends to “monster formation.”
There are a number of other small Stentor
species,
almost all of which are relatively easily cultured.

b2)
Bursaria
truncatella is
like an enormous (micro-world-wise) crystal Art Deco vase. It swims
through the water in a stately fashion and has an exceptionally
large, spiral cytostome which allows it to easily engulf Paramecia.
It’s a bit tricky to culture for more than a week or so. If
you want keep it going longer, you will need to keep it in a
relatively “clean” culture with sufficient food, but not
too much. If it has too little or too much in the way of food, it
tends either to encyst or disintegrate. It is essential to avoid
high levels of bacterial growth and frequent subculturing is
recommended.

b3)
Dileptus
anser
is a wonderful beastie whose anterior end, extending up from the
cytostome, waves through the water like the trunk of an elephant.
Its needs in culture are similar to those of Bursaria,
except that smaller food organisms, such as Colpidium,
rather than Paramecium,
should be provided.

b4)
Another of the giants of the protozoan world is Spirostomum ambiguum,
a long, band-shaped, highly contractile ciliate. Different species
of Spirostomum vary widely in size, but S. ambiguum is well over
1,000 microns when extended and can get up to 3,000 microns. When
disturbed, it does its magic act and contracts to about 1/3 its
previous length. It is also of interest because of its long chain
nucleus and an immense posterior contractile vacuole. I have
described Spirostomum as micro-whales, since they are the leviathans
of the micro-world and feed largely on bacteria which is parallel to
many of the great cetaceans feeding on plankton.

I
have kept long-term cultures (several years) of 3 different species:
the giant S.
ambiguum,
S.
minus which
is not quite so long and much narrower and with a shorter chain
nucleus, and S.
teres,
a rather small species (150 to 200 microns) with a single ovoid
nucleus. The matter of contractility alone makes them well worth
studying.

b5)
Blepharisma,
like Stentor
coeruleus,
has a distinctive pigment, only in this case, it is a pink to light
rose color. The pigment is unique and is called blepharismin. It is
a photoactive pigment and acts as a biochemical signal; if
Blepharisma
is
exposed to strong light for a prolonged period, the pigment becomes
toxic to the organism and eventually lethal–something to
remember when you are examining this beastie for long periods under
the intense illumination of the microscope.

All
of these large ciliates culture reasonably well in Giese salts with
either a grain of wheat or rice added.

Other
ciliates have rather special dining habits and need to be “spoon-fed”
as it were. Didinium
nasutum is
probably the most common example and feeds almost exclusively on
Paramecia. I have read a number of accounts which inform me that
when the Paramecium
population
has seriously diminished that the Didinium
will
begin to form cysts and that all one has to do to revive a culture is
add a fresh supply of Paramecia–or that if you have let the
culture go until only Didinium
cysts
remain, that you can keep them in water, add Paramecia months later
and the Didinium
will
excyst and produce a flourishing culture. To this I emphatically
say–Nonsense! Over the years, and as recently as last month, I
have tried this at least a dozen times with no
success. I mention this, because sometimes, despite your best
efforts, an organism may frustrate your attempts while seemingly
everyone else has great success. A late colleague of mine, who was a
distinguished cell biologist and an exceptionally able and astute
protozoologist, asked me to provide him some specimens of Lacrymaria
olor,
which I did gladly–on quite a number of occasions. Despite the
fact that I had published a protocol for culturing this particular
organism and, that a fair number of people in other laboratories in
various places around the country had used it successfully, he was
never able to establish stable cultures in his lab. Secretly, I
always suspected that his graduate assistants were to blame. Here,
the message is, don’t get frustrated if you can’t succeed
with a particular organism. Micro-environments are exceedingly
complex. Today I was looking at a depleted Lacrymaria
culture
and discovered some bacterial colonies unlike any I had seen before.
Each had a center point from which emerged a balloon-like structure
and from the base a series of “spokes” radiating out like
a line of spaced beads which got smaller and smaller as their
distance from the center increased. So, that culture failed
Lacrymaria-wise,
but certainly succeeded in other respects.

c)
Large and medium-sized amoebae can be fairly readily subcultured and
kept thriving for months. It is advisable to use a salt solution
which you can make up in your own lab if you have the chemicals and a
good balance with a sensitivity to 0.01 gm. Alternatively, you can
buy amoeba medium from a biological supply house. In any case, you
can add a grain or two of wheat or rice or a few pieces of boiled
hay. When I have time I like to use all three in separate dishes.
Right now I have 4 subcultures of Amoeba
proteus going
and one of them (a rice/Giese salt culture) I have intentionally
rather neglected. Amoeba
proteus is
a magnificently strange organism and when neglected manifests an
astonishing rage of intriguing morphological variations. This
particular culture is certainly flourishing in terms of sheer
numbers, for there are hundreds of specimens in my little 2 inch
dish. Some look just like you would expect any self-respecting
Amoeba
proteus to
look, whereas others are hardly recognizable. Some have been feeding
in a irregular manner and appear quite black at 60x under my stereo
microscope. Others have ceased extending pseudopodia altogether and
have contracted into a spherical blob containing an enormous vacuole.
Yet others are typically extending just one long pseudopodium which
is atypical behavior for Amoeba proteus, but usual for a mature
Pelomyxa. I would like to photograph or videotape these many
morphological variations, but again I remind you that amoebae are
(especially large ones) dramatically 3-dimensional–put them
under a cover
glass and there’s a good chance that you’ll crush most,
if not all of them. So, what I hope to do is start a culture in a
special chamber for my inverted microscope, so that I can observe
and, it is hoped, photograph them without having to remove them from
the environment of their culture.

d)
Small amoebae and heliozoa sometimes show up in enormous numbers in
cultures of other organisms. This makes them fairly easy to
subculture if you get interested in studying them. They have some
very unusual characteristics and one can spend a great deal of time
investigating both their morphology and their behavior.

e)
Medium-size ciliates. Very small ciliates often move exceptionally
rapidly, which along with their size, makes them very difficult to
study, but there are a number of interesting ciliates of intermediate
size which are readily cultured and can be maintained in subculture
with not much effort. Many of them do quite nicely in rather rank,
neglected cultures. Hypotrichs thrive in a variety of conditions and
are thus quite adaptable. They are of special interest because of
their special organelles called “cirri” which are fused
bundles of cilia creating a thicker, stouter structure than the
traditional cilium. They often use their cirri to walk or scamper
across the bottom while feeding.

Another
ciliate which shows up very frequently in my old samples and cultures
is Cyclidium
glaucoma.
This organism has long caudal cilia rather like bristles and an
undulating membrane around the cytostome. Its behavior is both
fascinating and frustrating. It will sit motionless on the
substrate, sometimes for minutes at a time and then just at the
moment when you rotate you turret to examine it with a higher powered
objective, it bounces off out of the field of view as though it were
spring loaded. Halteria
grandinella is
also a leaper.

Coleps
is
at times quite common and will attack injured larger organisms such
as Paramecia. They are barrel-shaped, some have 4 or 8 spines toward
the posterior end, and they are armor-plated.

Colpoda
can
almost always be found by taking a sample of good soil, add a salt
solution, and a grain of wheat. These organisms are roughly
kidney-bean shaped and are voracious feeders on bacteria and minute
protists.

The
aptly named Urocentrum
turbo whizzes
through the field of view with such speed that only the most indolent
observer can ignore it. Then suddenly one will come to a screeching
halt, attach to the substrate with a bit of “glue” from
its tuft of longish caudal cilia and proceed to spin like a top. It
is marvelous to behold.

f)
Flagellates. Personally, I find trying to culture the more
interesting flagellates rather frustrating. Balanced salt media are
important along with low bacterial growth, plus a proper balance of
light, temperature, and oxygen. Volvox
are among the most lovely and astonishing denizens of the
micro-world; they look like great crystal spheres rolling through the
water and when they occur in great numbers, they tint the water a
delicate shade of green. They consist of hundreds of cells and
frequently have subcolonies growing in them. There is an intricate
lattice of connection between the cells which is clearly visible at
high magnifications. They obtain nutrition photosynthetically, thus
the green tint. In general, I have not had good luck with trying to
culture, let alone subculture, chlorophyll-bearing flagellates.
However, I have been able to maintain phytoflagellates for periods of
2 to 3 week and this provides sufficient time to do some intensive
examination. Smaller colonial flagellates, such as, Pandorina,
Gonium,
and Synura
are
certainly worth attention as well.

There
are a number of interesting species of “armored”
flagellates or dinoflagellates in fresh water, but the marine species
are of special concern. Sometimes both in lakes and ponds as well as
the ocean, algal “blooms” will occur and a few species
will show up in uncountable numbers. These are most serious in
marine coastal environments. Since certain dinoflagellates contain a
powerful toxin when shellfish and other filter feeders ingest them,
they can become toxic, even fatal, to humans who eat them. The
dinoflagellates which occur in the lakes and ponds here in the high
plains and mountains seem to prefer quite cold water and thus are not
easily cultured.

Euglenoids
are somewhat easier to culture and a medium made from split peas is
frequently used for them.

Two
brief sets of final remarks. Whenever you find an organism that’s
of interest and which is thriving–subculture it! I am all too
aware of the fact that this can quickly get out of hand without
having a separate room for maintaining cultures with a variety of
climate control features and a laboratory technician to oversee all
of this, preferably one who will do all of this work on a volunteer
basis because he or she is dedicated to science rather than that
awful stuff we call money. However, until Microsoft establishes the
Foundation for the Encouragement of Amateur Microscopy and offers
grants to supply us with equipment and technicians, we shall just
have to muddle along as best we can and if one looks at Micscape,
I think we have proved that indeed we can muddle along, but a bit of
financial encouragement would certainly be welcome. Perhaps we need
to hire some creative accountants.

My
absolute final remarks here have to do with a critical issue which I
haven’t yet broached–glassware (and plasticware). Never
use
glassware which has contained potential toxic chemicals either for
culturing or for the preparation of culture media. Glass is an
extraordinary substance and, in spite of its hardness and toughness,
it is easily contaminated at the micro-level. Anyone who employs any
fixatives using mercury salts, such as Schaudinn’s fluid,
should use disposable glass- and plasticware. Only amateurs who have
extensive experience in handling such dangerous chemicals should use
such fixatives at all. Not only will mercury salts contaminate
glassware, they are extremely
poisonous.
So, whatever chemicals you use in your lab, keep all glassware used
for reagents completely separate from dishes, beakers, flasks, or
pipets used for culture or the preparation of culture media. Soaps,
detergents, and even alcohols can leave fine films or deposits and
some of these may have adverse affects on the organisms you are
trying to culture. Cleanliness is next to godliness—but not always.
I would rather risk a few organic contaminants from old cultures than
try to figure out whether a soap, detergent or alcohol is creating a
problem with my culottes.

So,
my procedure is not very scientific, rather casual, and a bit sloppy,
but is has worked rather well for me for quite some years. I have
only a small bathroom sink for cleaning my culture dishes, so when I
get enough to fill the basin, I take a small plastic knife of the
sort one takes along on a picnic and I scrape out any excess
detritus, dried wheat or rice grains. I fill the basin with very hot
water and let the dishes soak for 1 or 2 hours. After that, a
medium-sized, natural bristle brush can be used to do a final
cleaning of the inside of the dishes. The brush should be well
rinsed and not used for any other purpose. There are 3 drying
possibilities which I use. When I have a bit of space and things
aren’t too cluttered (which is very rare), I simply spread the
dishes out and let them air dry. Sometimes I dry them with paper
towels, preferably cheap, plain brown ones that haven’t been
treated with bleaches or dioxin or I will use a soft kitchen towel of
the sort that you used to use for drying dishes before you got your
dishwasher.

Several
years ago, I came across a super sale of plastic bottles . (I’m
a total sucker for sales. One of my friends claims that I would buy
typewriter ribbons if they were on super sale, even though I no
longer own a typewriter. I think that’s a bit of an
exaggeration, but I do admit to having made some strange purchases.)
The bottles were 8 ounces and had plastic screw cap lids–quite
suitable for collecting pond samples, so I bought 50 of them. These
were 20 cents each, so for the modest investment of 10 dollars, I
have enough to last me the rest of my life, since I use and reuse
them. I also never wash them. When I’m finished with a
sample, I either let it dry up or if it’s a smelly one, empty
it out in they alley and then let it dry in a box which I keep in the
garage for sample bottles. The next time I go collecting, I take a
twig and scrape out any excess, encrusting debris, fill the jar with
water, shake it vigorously, empty it and collect a new sample. If
there are some leftover cysts that insist on excysting after such a
long period of encystment, well–so be it.

A
Small Gallery of Protists

1.
Spirogyra.
A
common and lovely filamentous alga.

2.
Amoeba
proteus.
Perhaps the most famous of all amoebae and yet it is not that common
in samples. Fortunately, it is easily cultured and can be obtained
from many biological supply houses. This specimen came from a
reservoir about 25 miles east of Laramie.

3.
Colpidium
sp. This
is also a very common organism and cultures readily. It is important
because it can be used as a food organism for many other larger
ciliates. The color here is a consequence of Nomarski differential
interference contrast.

4.
Didinium
nasutum.
One of the most voracious ciliates. These images were taken from a
prepared slide. This first image is of a single Didinium up close
showing the distinct macronucleus, the two distinctive ciliary bands
which run around its circumference, and the prominent proboscis which
contains a series of structures called pharyngeal rods. These rods
allow the cytostome (mouth) to expand to a remarkable degree.

5.
Didinium
attacking
Paramecium.
Here we see Didinium
ferociously
attacking a Paramecium
and
as you can see the mouth has partially expanded.

6.
Didinium
devouring
Paramecium.
As you can see, Didinium
is
able to expand its mouth to such an extent that in can engulf an
entire Paramecium.

7.
Dileptus
anser.
This bizarre ciliate waves its “trunk” rather like an
elephant and at the base of the trunk is the cytostome which has a
grouping of toxicysts which it can discharge to stun potential prey.

8.
Spirostomum
minus.
This is one of the species of Spirostomum
which
I have elsewhere described as “micro-whales”. This
specimen was approximately 1200 microns in length. It is highly
contractile and has a beaded macronucleus.

9.
Spirostomum
minus.
I have stained this specimen causing it to contract and the stain
(Methyl Green Acetic) shows the beaded macronucleus clearly.

10.
Vorticella sp. Like Spirostomum
and
Stentor,
Vorticella
is
contractile. This lovely bell-shaped organism was first described by
Leeuwenhoek. The bell itself is contractile and there is a long
fiber called a myoneme running through the stalk, as you can see in
the image. When disturbed, the organism contracts and the stalk
coils up, then slowly uncoils later.

11.
Stentor
coeruleus.
This species has a distinctive dichroic pigment which usually gives
it a blue-green appearance, however, when the light shifts to just
the right angle, the organism takes on a rose-colored tint. This
specimen was swimming and is not fully extended.

12.
Paramecium.
Along with Amoeba
proteus,
perhaps the most famous of all the protozoans, Paramecium is anything
but a typical ciliate. It has many fascinating morphological
features, is easy to culture, and is an excellent organism for
conducting a wide variety of experiments.

13.
Thecamoeba.
These remarkable creatures have a textured membrane, are often found
in cultures made from soil taken from potted plants, and have both
unusual morphological and behavioral characteristics. You can see
that it has ingested a rotifer and some filamentous algae.

14.
Thecamoeba. This specimen was taken from a dish to which I added a
weak solution of the stain Neutral Red. It was sufficiently dilute
that it did not kill the Thecamoeba,
but it did give a vivid color to the filamentous algae which the
organism then later ingested. This is a helpful way to study the
feeding behavior of certain micro-organisms.